ion exchangers nhx1 and nhx2 mediate active potassium ... · ion exchangers nhx1 and nhx2 mediate...

Ion Exchangers NHX1 and NHX2 Mediate Active PotassiumUptake into Vacuoles to Regulate Cell Turgor and StomatalFunction in Arabidopsis W OA

Veronica Barragan,a Eduardo O. Leidi,a Zaida Andres,a Lourdes Rubio,b Anna De Luca,a

Jose A. Fernandez,b Beatriz Cubero,a and Jose M. Pardoa,1

a Instituto de Recursos Naturales y Agrobiologia, Consejo Superior de Investigaciones Cientificas, Sevilla 41012, Spainb Departamento de Biologia Vegetal, Facultad de Ciencias, Universidad de Malaga, Malaga 29071, Spain

Intracellular NHX proteins are Na+,K+/H+ antiporters involved in K+ homeostasis, endosomal pH regulation, and salt

tolerance. Proteins NHX1 and NHX2 are the two major tonoplast-localized NHX isoforms. Here, we show that NHX1 and

NHX2 have similar expression patterns and identical biochemical activity, and together they account for a significant

amount of the Na+,K+/H+ antiport activity in tonoplast vesicles. Reverse genetics showed functional redundancy of NHX1

and NHX2 genes. Growth of the double mutant nhx1 nhx2 was severely impaired, and plants were extremely sensitive to

external K+. By contrast, nhx1 nhx2 mutants showed similar sensitivity to salinity stress and even greater rates of Na+

sequestration than the wild type. Double mutants had reduced ability to create the vacuolar K+ pool, which in turn provoked

greater K+ retention in the cytosol, impaired osmoregulation, and compromised turgor generation for cell expansion. Genes

NHX1 and NHX2 were highly expressed in guard cells, and stomatal function was defective in mutant plants, further

compromising their ability to regulate water relations. Together, these results show that tonoplast-localized NHX proteins

are essential for active K+ uptake at the tonoplast, for turgor regulation, and for stomatal function.

INTRODUCTION

Potassium (K+) is an essential macronutrient that fulfills important

functions related to enzyme activation, osmotic adjustment and

turgor generation, regulation of membrane electric potential, and

cytoplasmic pH homeostasis. K+ is acquired by roots, redistrib-

uted among plant tissues and organs, and stored in large

quantities inside vacuoles, and it is the most abundant inorganic

cation in plants, comprising up to 10% of their dry weight (White

and Karley, 2010). Most terrestrial plants are able to grow in a

wide range of external K+ concentrations, from lowmicromolar to

10 to 20 mM levels (Rodrıguez-Navarro, 2000). K+ uptake by

plant roots is thought to be facilitated by two independent

transport mechanisms with distinct kinetic parameters and se-

lectivity (Epstein et al., 1963). The high-affinity, K+ selective, and

saturable System 1 operates in the micromolar range andmoves

K+ into the cytosol of root cells against the electrochemical

gradient. Electrophysiological evidence indicates that this path-

way involves a H+:K+ symporter coupled to the activity of the

plasma membrane H+-ATPase and is capable of driving K+

accumulation ratios in excess of 106-fold (Maathuis and

Sanders, 1994). Molecular genetic approaches have implicated

high-affinity K+ uptake permease (HAK/KUP) transporters in this

process (Santa-Marıa et al., 1997; Gierth et al., 2005; Rodrıguez-

Navarro andRubio, 2006). The low-affinity pathway, or System 2,

has the characteristics of channel-mediated transport and dom-

inates K+ uptake at external K+ concentrations of above 0.5 to

1 mM and usual plasma membrane potentials of 2120 to 2200

mV. Channels of the Shaker family have been implicated in this

passive K+ permeation through the plasma membrane downhill

the K+ electrochemical gradient. In Arabidopsis thaliana, the

voltage-gated K+-selective channel protein K+ Transporter1

(AKT1) has been shown to participate in K+ uptake by roots

(Hirsch et al., 1998; Spalding et al., 1999; Xu et al., 2006). Stelar

K+ Outward Rectifier is structurally similar to AKT1 but mediates

K+ efflux in root stellar cells to facilitate K+ loading into the xylem

(Gaymard et al., 1998). InArabidopsis guard cells, wheremassive

K+ fluxesmediate stomatal movements, stomatal opening driven

by K+ entry occurs mainly through K+ channel in Arabidopsis

thaliana 1 (KAT1) and KAT2, whereas stomatal closing is caused

by K+ efflux through Gated Outwardly Rectifying K+, a channel

activated by membrane depolarization (Ache et al., 2000; Kwak

et al., 2001; Hosy et al., 2003; Lebaudy et al., 2010).

Compared with the plasma membrane, much less is known

regarding K+ transport processes at the vacuole, although in-

sights into the transport mechanisms and proteins involved in K+

fluxes across the tonoplast are now emerging. Two different

proton pumps energize the tonoplast: the V-ATPase that is

powered by ATP and the tonoplast-bound pyrophosphatase that

hydrolyzes inorganic pyrophosphate (V-PPase). In most species

and cell types, both H+ pumps generate pH gradients of 1 to 2 pH

units (acidic inside) and an electrical charge (membrane

1Address correspondence to [email protected] author responsible for distribution of materials integral to thefindings presented in this article in accordance with the policy describedin the Instructions for Authors (www.plantcell.org) is: Jose M. Pardo([email protected]).WOnline version contains Web-only data.OAOpen Access articles can be viewed online without a subscription.www.plantcell.org/cgi/doi/10.1105/tpc.111.095273

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potential) of 20 to 40 mV that is positive in the vacuolar lumen

relative to the cytosol. This fact implies that positively charged K+

ions are excluded from the vacuole in K+-replete cells unless

transport is coupled to an energy-dependent uptake mecha-

nism, whereas efflux could be driven by channels that permeate

K+ downhill its electrochemical gradient. Electrophysiological

and genetic evidence has shown that upon a decrease in

cytoplasmic K+, slow-activating vacuolar (SV) channels release

K+ from vacuoles to assist in the homeostatic partition of K+

between the cytoplasmic and vacuolar pools (reviewed in

Isayenkov et al., 2010; Hedrich and Marten, 2011). Several

HAK/KUP transporters (Arabidopsis KUP4, KUP5, and KUP7)

have also been found in the tonoplast, and theymight catalyze K+

efflux from the vacuole below the electrochemical limits of

passive permeation by channels; however, functional evidence

is still lacking (Banuelos et al., 2002; Jaquinod et al., 2007).

Since transport of K+ into the vacuole in K+-replete cells proceeds

against theK+electrochemical gradient, aK+/H+antiporter energized

by the pH gradient across the tonoplast was thought to achieve

vacuolar K+ accumulation (Walker et al., 1996; Carden et al., 2003).

Scattered evidence indicated that vacuolar Na+,K+/H+ antiporter

(NHX)–type exchangers could serve this critical function in plant cells

(Venema et al., 2002; Rodrıguez-Rosales et al., 2008, 2009; Yoshida

et al., 2009; Jiang et al., 2010; Leidi et al., 2010). Studies in transgenic

tomato (Solanum lycopersicum) showed that overexpression of the

vacuolarAtNHX1or theendosomal LeNHX2Na+,K+/H+exchangers

led to greater K+ accumulation (Rodrıguez-Rosales et al., 2008; Leidi

et al., 2010). Recently, a reverse genetics approach in Arabidopsis

has shown that NHX1 and NHX2 proteins sustain the intravacuolar

K+ concentration and, in the process, regulate the vacuolar pH and

facilitate cell expansion (Bassil et al., 2011b).

Because vacuoles occupy most of the intracellular volume in

many plant cells and are the main cellular reservoir for K+,

changes in tissue K+ concentration are largely a reflection of the

dynamics of the vacuolar pool. By contrast, cytosolic K+ con-

centrations are tightly regulated through the integrated regula-

tion of K+ uptake and efflux at the plasma membrane and K+

import and export at the tonoplast (Leigh, 2001; White and

Karley, 2010). Cytosolic K+ concentration is thought to decline

only when the vacuolar K+ reserve has been depleted below the

thermodynamical equilibrium with the cytosolic pool (Walker

et al., 1996). Conversely, surplus K+ is placed into the vacuole to

maintain cytosolic K+ within narrow limits independently of K+

abundance in the growth medium. In K+-sufficient plants, the

vacuolar pool plays a chief biophysical function, the lowering of

osmotic potential to generate turgor and drive cell expansion.

Rapid cell expansion relies on high mobility of the active

osmoticum, and the highly permeant and abundant inorganic

ion K+ fits this role. There are numerous examples of K+ fluxes

driving cell expansion and organ movements (White and Karley,

2010), but nowhere is this critical function of the vacuolar K+ pool

more apparent than in guard cells. The rapid accumulation and

loss of K+ and anionic organic acids by guard cells, mostly in the

vacuolar compartment, regulates the opening and closing of

stomata and, thereby, gas exchange and transpiration. Stomatal

opening starts with membrane hyperpolarization caused by H+-

ATPases, which induces K+ uptake through inward-rectifying

K+in channels (reviewed in Kim et al., 2010). Influx of K+, Cl2, and

NO32 and the synthesis of malate increase turgor in the guard

cell and induce stomatal opening. The rapid loss of K+ fromguard

cells during stomatal closure is started by anion release via anion

channels, which causes plasma membrane depolarization and

the opening of K+ channels that facilitate K+ efflux from the

vacuole and across the plasma membrane. Among the solutes

released from guard cells, >90% originate from vacuoles.

Here,we report that the twomajor isoformsof vacuolar-localized

NHX proteins in Arabidopsis, NHX1 and NHX2, play a critical and

redundant role in the accrual of K+ in the vacuole, which in turn

impinges on the ability of plants to generate turgor and sustain

osmotic regulation. Furthermore, we show that NHX1 and NHX2

genes are greatly expressed in guard cells of stomata, where they

contribute to the regulation of stomatal function and transpiration.

RESULTS

Biochemical Similarities of NHX1 and NHX2 Exchangers

Six genes (NHX1 toNHX6) encoding NHX exchangers have been

identified in the genomic sequence of Arabidopsis (Yokoi et al.,

2002). RNA gel blotting and RT-PCR, used to determine the

relative abundance of the NHX transcripts, showed that NHX1

andNHX2 transcripts were both abundant andwidely distributed

in plant tissues and that they accumulated in response to the

same effectors (NaCl, KCl, LiCl, osmotic stress, and abscisic

acid) (Yokoi et al., 2002; Aharon et al., 2003). At the protein level,

NHX1 and NHX2 are the most closely related members of the

NHX family in Arabidopsis (87.5% sequence identity) and are

both localized in the tonoplast (Yokoi et al., 2002; Bassil et al.,

2011b). NHX1 has been shown to mediate Na+/H+ and K+/H+

exchange with similar affinity (Venema et al., 2002; Yamaguchi

et al., 2005; Hernandez et al., 2009). To test whether NHX2 had

similar transport properties as NHX1, NHX2 tagged with a hexa-

His tag at the C terminus was expressed in budding yeast

(Saccharomyces cerevisiae) cells and purified by Ni2+ affinity

chromatography as previously described for NHX1 (Venema

et al., 2002). Phenotypic complementation of a mutant nhx1

yeast strain showed that tagged NHX2:H6 was functional and

indistinguishable from the untagged, wild-type protein (Figure

1A). The NHX2:H6 protein was purified and inserted in soybean

(Glycine max) phospholipid vesicles formed in reconstitution

buffer containing the pH indicator pyranine and (NH4)2SO4.

Dilution of the proteoliposomes in reconstitution buffer without

(NH4)2SO4 produced an instantaneous diminution of pyranine

fluorescence due to leakage of NH3 and the subsequent internal

acidification of the vesicles (Venema et al., 2002). Cation/proton

exchange was initiated upon the addition of NaCl or KCl salts,

and the rate of pH variation inside the vesicles was estimated

from the change in pyranine fluorescence (Figure 1B). NaCl and

KCl salts produced similar rates of fluorescence recovery.

Replacing chloride salts with gluconate, a relatively impermeable

anion, produced similar fluorescence recovery rates, demon-

strating that cations were exchanged for intravesicular protons

(Figure 1D). The small increase in net exchange with gluconate

salts relative to chloride salts indicated that only a small fraction

of protons that were transported out of the vesicles in the

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exchange reaction leaked back into the vesicles together with

chloride anions. Importantly, mixing equal amounts of sodium

and K+ gluconate salts produced the same exchange rate as

either salt alone, demonstrating that NHX2 lacked significant

discrimination between Na+ and K+. Exchange reactions cata-

lyzed by NHX2 displayed saturation kinetics (Figure 1C) and the

Km estimated for Na+ and K+ ions was ;35 mM (R2 = 0.999).

Similar results have been obtained with NHX1 (Venema et al.,

2002; Hernandez et al., 2009; Leidi et al., 2010). Therefore, NHX1

and NHX2 appear to be biochemically equivalent Na+,K+/H+

exchangers.

Expression Pattern of NHX2

We next compared the gene expression pattern of NHX2 to that

of NHX1. The latter gene has been shown to be expressed in

nearly all tissues throughout plant development (Shi and Zhu,

2002; Apse et al., 2003). To determine the tissue expression

pattern of NHX2, an ;3.1-kb promoter region upstream of the

NHX2 start codon was fused with the b-glucuronidase (GUS)

reporter gene, and the resulting construct, ProNHX2:GUS, was

transformed into wild-type Arabidopsis plants. Three indepen-

dent transgenic lines were assayed for GUS expression and they

produced consistent patterns. GUS expression was detected at

all developmental stages tested, from seed germination to

flowering and seed setting. As depicted in Figure 2, histochem-

ical GUS staining was detected, with varying intensity, in most

tissues of Arabidopsis seedlings. In leaves and hypocotyls, GUS

staining was stronger around the vasculature, in meristems, and

in guard cells of the stomata (Figures 2A to 2D). In roots, the

strongest GUS activity was detected in the vasculature, at the

root tip, and at the points of emergence of secondary roots, but

not in root hairs and the epidermis (Figures 2D to 2F). In flowers,

GUS staining was restricted to the filament of the stamens, the

ovarian stigma, mature pollen grains within the anthers, and the

pollen tube (Figures 2G to 2K). Muchweaker staining was seen in

sepals, and it was associated with vascular tissues. No signifi-

cant GUS activity was detected in petals (Figure 2G). Remark-

ably, the strongest GUS staining in flowers was also observed in

stomata, including those in the anthers (Figure 2J). In immature

siliques, GUS staining was restricted to the septum and to the

silique tip and base (Figure 2L). This expression pattern strongly

resembled that of NHX1 (Shi and Zhu, 2002), except at the tip of

main roots, root hairs, and meristems, in which NHX1 and NHX2

promoter constructs showed opposite expression levels. Public

microarray databases also indicate similar expression patterns

of NHX1 and NHX2 genes and the significant greater abundance

of their transcripts in guard cells of Arabidopsis (Winter et al.,

2007). However, the various divergences regarding their expres-

sion patterns imply extensive overlapping, but not identical

physiological functions of NHX1 and NHX2. The tissue expres-

sion pattern of NHX2 was also examined in seedlings after

sodium, lithium, and sorbitol treatments that were reported to

induce the accumulation of NHX2mRNA (Yokoi et al., 2002), but

no significant deviation from the pattern found under control

conditions was observed, suggesting that stress environments

upregulated expression without altering tissue specificity.

Functional Redundancy of NHX1 and NHX2

Previous work showed that NHX1 and NHX2 are functionally

redundant and that nhx1 and nhx2 singlemutants havemoderate

disturbances in the germination rate, biomass production, and

foliar area compared with wild-type plants (Apse et al., 2003;

Bassil et al., 2011b). This is in agreement with the extensive

overlapping expression pattern ofNHX1 andNHX2 (Figure 2) (Shi

and Zhu, 2002), identical subcellular localization of the proteins

(Yokoi et al., 2002; Bassil et al., 2011b), and similar transport

activity (Figure 1) (Venema et al., 2002). To investigate further the

physiological role of vacuolar NHX proteins, we sought to

Figure 1. Biochemical Activity of NHX2.

(A) Complementation of the yeast nhx1mutant (top), transformed with an

empty vector, or to express NHX1, NHX2, or His-tagged NHX2 (four

independent transformants), in Arg phosphate medium supplemented

with 70 mM NaCl and yeast peptone dextrose (YPD) medium with 30 mg/

mL hygromycin B (HygB).

(B) Typical exchange reaction in proteoliposomes. Arrows indicate the

addition of substrate salts NaCl or KCl (1). SO4(NH4)2 was added to

collapse the proton gradient and end the assay (2). Pyranine fluores-

cence at 510 nm is given in arbitrary units.

(C) Saturation kinetics of exchange rates catalyzed by NHX2 as a

function of substrate (NaCl) concentration.

(D) Exchange rates using as substrate 45 mM sodium (NaGlu) or K+ (KGlu)

gluconate salts, 22.5 mM each of these salts (Na/K), and 45 mM NaCl or

KCl. Shown is the mean exchange rates of two independent assays.

NHXs Regulate Plant Turgor and Stomata 1129

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produce homozygous null double mutants with alleles nhx1-2

and nhx2-1 carryingmutationswithin the coding regions ofNHX1

and NHX2, respectively (Bassil et al., 2011b) (see Supplemental

Figure 1A online). However, early attempts to produce double

knockout plants with nhx1-2 nhx2-1 null alleles were unsuccess-

ful. To circumvent this problem, we used another allele, nhx1-1,

that carries the T-DNA insertion in the 59-untranslated region of

NHX1 (at nucleotide 2114; see Supplemental Figure 1A online).

Contrary to mutant nhx1-2, which failed to produce detectable

levels of gene transcripts as determined by RNA gel blot and RT-

PCRassays,mutantnhx1-1 showeda low, yet detectable, amount

of NHX1 transcript, which accumulated slightly under salt stress

(see Supplemental Figure 1B online). Mutant nhx2-1 produced a

59-truncated mRNA that accumulated at greater levels than in

wild-type plants (see Supplemental Figure 1C online).

Homozygous lines of genotype nhx1-1 (leaky allele) and

nhx2-1 were crossed, and homozygous double mutants were

identified by diagnostic PCR amplification of NHX1 and NHX2.

Only two homozygous nhx1-1 nhx2-1 plants were found in the F2

population (n = 72), and these plants were severely stunted and

failed to produce seeds. To increase the chances of isolating a

larger number of double homozygous nhx1-1 nhx2-1mutants, an

F2 plant of genotype nhx1-1/nhx1-1 NHX2/nhx2-1 was self-

pollinated, and homozygous mutants nhx1-1 nhx2-1 were re-

covered among the F3 progeny, albeit with a frequency lower

than expected (7.7% instead of 25%, two out of 26 plants

genotyped by diagnostic PCR), further indicating that not only

the vegetative growth of double nhx1-1 nhx2-1 mutants was

compromised, but also their viability. Two lines of genotype

nhx1-1 nhx2-1 that produced seeds, L2 and L14, were selected

for further study. Note that lines L2 and L14 share the same

genotype and that they should be regarded as biological repli-

cates. The absence of full-length NHX2 transcripts and the

residual levels of NHX1 mRNA in these lines were confirmed by

RT-PCR.

Amodified Long Ashtonmineral nutrient solution with 1mMK+

and nominally free of Na+ and NH4+ (LAK medium) was designed

to test the effects of K+ and Na+ on the growth of nhx1 nhx2

mutants. Avoidance of the high concentration of NH4+ in the

Murashige and Skoog medium routinely used for Arabidopsis

growth was important to prevent inhibition of K+ uptake at low

external K+ concentrations (Spalding et al., 1999; Rubio et al.,

2008). As recently described by Bassil et al. (2011b), plants of the

nhx1-1 nhx2-1 genotype were smaller than the wild-type and

Figure 2. NHX2 Promoter-GUS Expression Pattern in Transgenic Arabidopsis Plants.

(A) GUS activity detected in cotyledons.

(B) Strong GUS staining in guard cells of stomata in mature leaves.

(C) to (E) Preferential expression in the vasculature of the main stem (C) and roots (E). Note the strong expression in the root-shoot transition (D) and the

point of emergence of secondary roots (E).

(F) Strong GUS staining in the root meristems.

(G) to (L) Expression in reproductive organs was greater in filaments of the stamen ([G] and [I]), stigma ([G] and [H]), mature grain pollen (I), and silique

septum (L). Note the high expression in the stomata of anthers in (J), which is a close-up image of (I), and in the elongating pollen tube (marked by arrow) (K).

Bars = 20 mm in (K), 50 mm in (B), 100 mm in (A), (C), to (F), and (I), and 400 mm in (G) and (L).

1130 The Plant Cell

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single mutant plants when grown in soil under standard growth

conditions and in hydroponic culture in LAK medium (see Sup-

plemental Figure 2 online). Importantly, nhx1-1 nhx2-1 plants

were prone to wilting under our regular growth conditions (day/

night settings: 25/206 28C, 40/60% relative humidity, 14 h light),

suggesting that failure to obtain double knockout nhx1-2 nhx2-1

plants might have been due to severely damaged water relations

in the complete absence of NHX1 andNHX2 proteins. The strong

expression of ProNHX2:GUS (Figure 2) and ProNHX1:GUS (Shi

and Zhu, 2002) promoter fusions in guard cells lent further

support to this hypothesis. Therefore, double mutant plants

carrying the null alleles nhx1-2 and nhx2-1 were produced by

self-pollination of a plant of genotype nhx1-2/nhx1-2 NHX2/

nhx2-1 and homozygous nhx1-2 nhx2-1 plants were recovered

under high relative humidity (above 60%). In these conditions,

homozygous mutants nhx1-2 nhx2-1 were obtained with one-

quarter frequency among the progeny (16 out of 44 plants, 3:1

segregation ratio with P < 0.05, x2 test). Knockout plants

displayed an extremely stunted growth, were highly sensitive

to humidity fluctuations, and produced small siliques with few

viable seeds (Figure 3). Because leaky mutants of genotype

nhx1-1 nhx2-1 had an array of phenotypic disorders similar to but

less extreme than those of the complete knockouts nhx1-2

nhx2-1, and they were easier to handle and propagate, subse-

quent experiments were done with the leaky lines L2 and L14

(nhx1-1 nhx2-1 genotype) except when indicated otherwise.

Reduced NHX Activity in the nhx1 nhx2 Double Mutant

To quantitate the accumulated contribution of NHX1 and NHX2

proteins to Na+,K+/H+ exchange in the vacuole, K+/H+ and Na+/

H+ antiporter activity was assayed in tonoplast vesicles from the

leaves of wild-type and nhx1-1 nhx2-1 plants (lines L2 and L14)

by following the relaxation of the pH gradient created by the

V-type H+-ATPase. Calculation of initial rates of fluorescence

recovery after the addition of KCl or NaCl showed that nhx1 nhx2

mutant lines L2 and L14 exhibited much lower Na+/H+ and K+/H+

exchange activity than wild-type plants (Figure 4A). At 50 mM

NaCl or KCl, the nhx1 nhx2 mutants retained less than one-third

the exchange activity of the wild type (Figure 4B), demonstrating

that NHX1 and NHX2 account for a large proportion of the total

monovalent cation exchange capacity of the tonoplast. Similar

results were obtainedwith tonoplast vesicles isolated from roots.

The commensurate reduction of exchange activity of Na+ and K+

ions is in agreement with the NHX activity previously determined

for Arabidopsis NHX1 (Venema et al., 2002; Hernandez et al.,

2009) and NHX2 herein (Figure 1).

NHX1 and NHX2 Are Essential to K+ Homeostasis

Growth of lines L2 and L14 nhx1 nhx2 mutant plants was

progressively reduced relative to Columbia-0 (Col-0) at increas-

ing K+ concentrations (from 0.1 to 20 mM K+) in hydroponic

culture (Figure 5; see Supplemental Table 1 online; note that

these two data sets belong to independent experiments). High

KCl produced a striking toxicity to nhx1 nhx2mutants that led to

leaf desiccation and eventual plant death (Figure 5). Toxicity was

specifically linked to K+ ions since 10 mM K2SO4 produced

effects similar to 20mMKCl. The knockout plants with null alleles

nhx1-2 nhx2-1 grew poorly in all K+ regimes in hydroponic

culture, but high KCl concentrations further aggravated growth

impairment (see Supplemental Figure 2 online). Genetic com-

plementation of line L14 with the cDNA of NHX2 under the 35S

promoter partially restored growth parameters under various KCl

regimes (see Supplemental Figure 2 online). In contrast with K+

ions, nhx1-1 nhx2-1mutant plants did not show greater suscep-

tibility to Na+ toxicity compared with the wild type (Figure 6).

Notably, nhx1 nhx2mutant plants accumulatedmore Na+ in their

shoots than the wild type at 50 to 100 mM NaCl (Figure 6; see

Supplemental Table 1 online).

K+ concentration analyzed in leaf sap extracts and expressed

on a tissue water basis (i.e., molar concentration) showed a

significantly greater K+ concentration in nhx1-1 nhx2-1 mutants

at high K+ in the hydroponic solution (Figure 5; see Supplemental

Table 1 online). However, lower K+ concentrations were found on

a dry weight basis (i.e., percentage of content) in shoots of nhx1

nhx2 double mutants in plants growing at various K+ concentra-

tions (Figure 5; see Supplemental Table 1 online). Significant

differences in shoot water content among lines explained this

paradox, as K+ supply affects the water content of plants (Leigh

and Wyn Jones, 1984). Plants of the nhx1 nhx2 genotype

consistently accumulated less water in their shoots than the

wild type, which translated into greater K+ mM concentrations,

even though they had a lower K+ content on a dry matter basis

(Figure 5; see Supplemental Table 1 online). In roots, where no

differences in water content among lines were found, K+ con-

centrationswere significantly lower in the doublemutants except

at 1mMK+ (seeSupplemental Table 1 online). Similar resultswere

obtained with a complete knockout line of genotype nhx1-2

nhx2-1 (seeSupplemental Figure 2 online), albeit growth reduction

Figure 3. Impaired Development of Double Null nhx1 nhx2 Mutant.

Depicted are inflorescences and rosettes of double knockout (KO)

mutant plants of genotype nhx1-2 nhx2-1 and of wild-type Col-0 plants

grown in hydroponic LAK medium. Note the diminutive siliques in the

mutant plant.


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due to K+ toxicity was less evident than in the leaky nhx1-1

nhx2-1 lines because growth of nhx1-2 nhx2-1 plants was

extremely impaired even at the lowest K+ concentration tested.

At 20mMKCl, shoots of the knockout plants had only half the K+

and water contents of the wild type (see Supplemental Figure 2

online).

Together, these results demonstrate that NHX1 and NHX2 play

a major role in the accrual of K+ in shoot tissues of Arabidopsis,

whereas they appeared to be dispensable for Na+ compartmen-

tation.Comparedwith thewild type,Na+accumulationproceeded

to an even greater extent in the shoots of the nhx1 nhx2 mutant

upon exposure to salinity stress (Figure 6).

Reduced Vacuolar K+ Pool Affected Turgor,

Cell Expansion, and Stomatal Function

To estimate the size of the vacuolar K+ pool in the nhx1 nhx2

mutant, freeze-fractured leaves of L14 plants grown in LAK

medium with 1 mM K+ were analyzed with a scanning electron

microscope fitted with energy-dispersive x-ray spectroscopy

(EDX). This technique analyzes the elemental composition of

cells and cell compartments in plant tissues. The large vacuoles

of mesophyll palisade cells of mutant plants showed fewer K+

counts than Col-0 (Figure 7). K+ counts were also significantly

lower in guard cells of stomata of mutant plants, where both

NHX1 and NHX2 show high expression levels (Figure 2; Shi and

Zhu, 2002), as well as in epidermal subsidiary cells neighboring

the stomata.

The uptake of K+ by plant cells, and its accumulation in

vacuoles, is the primary driver for turgor generation and cell

expansion (Leigh, 2001). Plants of lines L2 and L14 showed

smaller leaf areas than Col-0 when grown at various K+ concen-

trations (see Supplemental Table 2 online). Specific leaf weight

and succulence were also significantly reduced in the nhx1-1

nhx2-1 mutant plants, suggesting that leaf expansion was hin-

dered by lower water uptake in the mutant (see Supplemental

Table 2 online). In freeze-fractured sections for scanning electron

microscopy, leaves from the nhx1-1 nhx2-1 leaky mutant (lines

L2 and L14) presented a less developed spongy parenchyma

when grown at 1 mM K+ (Figures 8A and 8B). At 10 mM K+,

palisade cells were significantly smaller compared with the wild

type and pyramidal in shape (Figures 8C and 8D). Smaller

mesophyll cells resulted in a thinner leaf lamina in the nhx1-1

nhx2-1mutant (Figures 8E and 8F), in agreement with the smaller

specific leaf weight (see Supplemental Table 2 online). Notably,

stomata of mutant lines showed aberrant morphology with

thinner guard cells compared with the wild type, indicating that

the lack of NHX1 and NHX2 function severely affected the

capacity of guard cells to expand (Figure 8G).

We next examined the ability of nhx1 nhx2 mutant plants to

withstand osmotic stress. Three-week-old plants of single mu-

tant lines (nhx1-2, nhx1-1, and nhx2-1) and double mutants

nhx1-1 nhx2-1 were transferred individually to 50-mL test tubes

adapted to hydroponic culture and filled with LAKmedium (1mM

K+) supplemented with 20% (w/v) polyethylene glycol 6000

(PEG6000) (Figure 9A). Plants were weighted at the time of

transfer and then at 0.5, 1, 2, 4, and 6 h afterward. Plants with

genotype nhx1-1 nhx2-1 suffered a significantly greater water

loss than the wild-type control (Figure 9B). Single mutant plants

experienced an intermediate water loss. After treatment with

PEG, wild-type and doublemutant plants had their roots washed

and were transferred to fresh LAK (1 mM K+) medium for

recovery. Wild-type plants flowered and produced healthy si-

liques, whereas the mutant plants accumulated anthocyanins,

their flower buds wilted, and they failed to produce siliques

(Figure 9C).

SinceNHX1 andNHX2were strongly expressed in guard cells,

we examined stomatal conductance in nhx1-1 nhx2-1 mutants

and its response to osmotic stress. The rates of transpiration and

photosynthesis were recorded with an infrared gas analyzer in

3-week-old plants of wild-type Col-0 and of mutant lines L2 and

L14 grown in hydroponic LAK medium. Without osmotic stress,

the stomatal conductance was significantly greater (P < 0.05;

n = 12) in the mutant lines (0.195 and 0.217 mol water m22 s21,

respectively) than in the wild type (0.123 mol water m22 s21).

However, the photosynthetic rateswere similar (Col-0, 5.33mmol

CO2 m22 s21; L2, 5.18 mmol CO2 m22 s21; L14, 5.01 mmol CO2

m22 s21). For stress treatment, leaves in the chamber were

allowed to equilibrate for at least 10 min and then plants were

Figure 4. Reduced Cation/Proton Exchange in nhx1 nhx2 Mutants.

(A) K+/H+ and Na+/H+ in tonoplast vesicles from leaves of Col-0 (black

circles) and mutant lines L2 (white circles) and L14 (white squares) over a

range of substrate cation concentrations.

(B) Average rates of cation exchange at 50 mM were significantly

reduced in the mutant line L14 (nhx1 nhx2) relative to Col-0.

1132 The Plant Cell

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subjected to osmotic shock by 20% PEG6000 (arrows in Figure

10). In response to the hyperosmotic treatment, stomata of wild-

type plants closed rapidly and reduced gas exchange by half in

;7 min (Figure 10A). By contrast, stomata from mutant plants

responded slowly, took 25 min to reduce transpiration to 50%,

and reached wild-type values only after 1.5 h. The slow response

ofmutant plants provoked the collapse of photosynthesis, whose

rate dropped to negligible values after 1.5 h (Figure 10B). The

experiment was repeated five more times with different mutant

and control plants, and similar results were obtained. These data

indicate that stomatal function and response to environmental

clues are impaired in plants lacking NHX1 and NHX2.

Stomatal action directly influences leaf turgor (Ache et al.,

2010). To inspect leaf turgor using intact Arabidopsis leaves, we

used a patch-clamp pressure probe to monitor turgor pressure

noninvasively. Continuous recording of leaf turgor showed sig-

nificant differences between the wild type and the nhx1-1 nhx2-1

mutant. Wild-type plants responded to the light stimulus with a

sudden decrease in leaf turgor followed by turgor recovery when

the light was off (Figure 11). These diurnal rhythms arise from

stomatal opening and closing, which regulate the rates of tran-

spiration (Ache et al., 2010). In plants growing in hydroponic

culture at low (0.1 mM) and medium (1 mM) K+ concentrations,

leaves from themutant line showed a steady loss of turgor during

the period of measurement. Moreover, turgor pressure oscilla-

tions in dark/light transitionswere erratic in themutant compared

with the wild type. After raising the K+ concentration in the

nutrient solution from 1 to 10 mM (arrow in Figure 11B), Col-0

plants showed a shift toward greater turgor pressures with

clearly marked dark/light oscillations, whereas the mutant

showed the opposite trend (i.e., moderate loss in average turgor

while dark/light oscillations were essentially absent). These

results are coherent with the inability of nhx1 nhx2mutant plants

to osmoregulate, a phenomenon that appears to be aggravated

by high K+, since transfer to 10 mM external K+ clamped leaf

turgor in the mutant and abrogated turgor oscillations in dark/

light transitions.

Cytosolic K+ Activity

Having established that nhx1 nhx2 mutant plants had a reduced

capacity to build up the vacuolar K+ pool, we next examined

whether cytosolic K+ was also affected in the mutant. To mea-

sure the cytosolic K+ activity (K+cyt) and membrane potentials

simultaneously, double-barreled K+-selective microelectrodes

were used to impale epidermal root cells of 15-d-old control and

Figure 5. Sensitivity of nhx1 nhx2 Mutant Plants to External K+.

(A) One-week-old seedlings of Col-0 and of mutant line L2 grown in LAK medium with 1 mM KCl were transferred to fresh hydroponic LAK medium

supplemented with the indicated concentrations of KCl. After 2 weeks, plants were collected and their shoot and root fresh weight were determined.

(B) Average fresh weight and SD values (n = 8 per line) of plants shown in (A). Differences between the mutant line and Col-0 were statistically significant

at P < 0.01 (a) or P < 0.05 (b) by the Fisher’s LSD method. Mutant line 14 produced identical results to L2.

(C) Average K+ and water content in shoots of Col-0 and nhx1 nhx2 mutant plants of line L2 (n = 6 per line) grown as in (A) in hydroponic LAK medium

with the indicated K+ concentrations. K+ content is given as a percent of dry weight (DW; top panel) and millimolar concentration (bottom panel). Water

content is given in the middle panel. Statistical differences by the LSD method of mutant relative to the wild type are indicated by letters; a, P < 0.01; b,

P < 0.05; c, P < 0.1. Line 14 produced identical results to L2.


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mutant plants of genotype nhx1-2 nhx2-1 grown in LAK medium

with 1 mM K+ (Figure 12). Cytosolic K+ activities were calculated

from calibration curves (slopes were close to 49 mV/p K+). The

average K+cyt was 756 14mMK+ (n = 5) in Col-0, which is similar

to previous estimates in root cells of wild-type Arabidopsis (8364 mM) (Maathuis and Sanders, 1993). In the nhx1 nhx2 mutant,

average K+cyt was 112 6 17 mM K+, which is significantly

different from wild-type values at P = 0.0086 (Student’s t test).

By contrast, membrane potentials were not significantly different

(P = 0.3123, Student’s t test) in thewild type (21226 14mV, n= 5)

and nhx1 nhx2 mutant (2132 6 14 mV, n = 5). These results

indicate that the impaired K+/H+ exchange at the tonoplast of

nhx1 nhx2 mutants elicited a significant increase of the cytosolic

K+ pool.

Decreases in cytosolic K+ content are thought to be required

for induction and development of high-affinity K+ uptake in

Arabidopsis (Rubio et al., 2008). Three-week-old plants of Col-0

andmutant lines L2 and L14 growing in LAK hydroponicmedium

with 1 mM K+ where starved in K+-free medium for 3 d and then

assayed for ion uptake rates by roots using rubidium (Rb+) at low

(100 mM) concentration as a tracer for K+. Short-term (#5 min)

uptake rates showed small differences among lines that were

not statistically significant, although roots from mutant plants

achieved a lower net uptake of Rb+ than the wild type over time.

Rb+ concentration in root tissues of Col-0 was 0.36 0.04 (mM6SD) after 20min, but only 0.186 0.02 in line L2 and 0.196 0.07 in

line L14. These results suggest that K+ uptake rates are some-

what reduced in themutant lines, presumably as a consequence

of greater K+cyt in the mutant.

DISCUSSION

The Essential Role of NHX Proteins in Creating

the Vacuolar K+ Pool

K+ is the major ionic osmoticum in plant cells and occurs in two

major pools, in the vacuole and in the cytosol. Cytosolic K+ plays

essential roles as activator of biochemical processes, in the

regulation of cytosolic pH, and in the fine-tuning of the plasma

membrane electrical potential. These fundamental functions

demand the maintenance of the cytosolic K+ concentration

within narrow limits (75 to 100 mM), regardless of changes in

K+ supply (Walker et al., 1996; Leigh, 2001). Since vacuolar K+

concentration closely follows K+ availability, the vacuolar pool is

the largest and most dynamic reservoir. In this compartment, K+

functions as an osmoticum to generate turgor and drive cell

expansion. Intracellular osmolytes reduce the cell water potential

and give rise to a passive water influx, which in turn increases cell

volume (Leigh and Wyn Jones, 1984). The lowest limit for

vacuolar K+ concentration appears to be 10 to 20 mM, which isFigure 6. nhx1 nhx2 Mutants Are Not Sensitive to Sodium.

Plants of Col-0 and of nhx1-1 nhx2-1 mutant line L2 (n = 4 per line) were

grown for 3 weeks in hydroponic LAK medium with 1 mM KCl and

supplemented or not with 50 and 100 mM NaCl. Shown are average and

SD values (n = 24)

(A) Fresh weight of shoots and roots of the wild type and mutant line L2.

(B) Sodium content in the shoot of plants after salinity treatment. DW, dry

weight.

Figure 7. Reduced Vacuolar K+ Content in the nhx1 nhx2 Mutant.

K+ content in the vacuoles of guard cells (GC), epidermal cells neigh-

boring the stomata (EC), and in mesophyll palisade cells (MP) of leaves

as determined by scanning electron microscopy/EDX. Shown are the

average percentage and SD of K+ counts relative to total elemental

counts. A minimum of 20 cells of each cell type per line was analyzed.

Statistical differences by the LSD method (P < 0.05) of mutant line L14

relative to the wild type are indicated by asterisks.

1134 The Plant Cell

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thought to reflect equilibrium with the cytosol at a maximum

trans-tonoplast voltage of ;40 to 60 mV (Leigh, 2001). Greater

concentrations of K+ inside the vacuole require active transport

from cytosol to vacuole that could be achieved by a nonelectro-

genic K+/H+ antiporter (Walker et al., 1996). Biochemical analy-

ses have shown that tonoplast-localized NHX proteins catalyze

the K+/H+ exchange with apparent affinity values (12 to 40 mM)

that are below the lower limits of cytosolic K+ concentrations and

are thus able to translocate K+ from the cytosol to the vacuole

under regular physiological conditions (Figure 1) (Venema et al.,

2002; Yamaguchi et al., 2005; Hernandez et al., 2009). Recently,

Figure 8. Impaired Leaf Cell Expansion in nhx1 nhx2 Mutants.

Freeze-fracture sections at scanning electron microscopy from leaves of

ArabidopsisCol-0 andmutant plants from line L14 (nhx1-1 nhx2-1) grown

in LAK medium for 2 weeks and then transferred to 1, 10, and 20 mM KCl

for an additional 2-week period. Equivalent leaves from each plant were

processed for scanning electron microscopy. Bars = 200 mM.

(A) Col-0 plants in 1 mM K+.

(B) L14 plants in 1 mM K+.

(C) Col-0 plants in 10 mM K+.

(D) L14 plants in 10 mM K+.

(E) Composition of serial pictures of a leaf from Col-0 plants grown for 2

weeks at 1 mMK+ and then transferred to 20 mMK+ for another 2 weeks.

(F) Composition of an equivalent leaf from a plant of mutant line L14

treated as in (E).

(G) Morphology of guard cells in stomata of wild-type, L2, and L14

plants.

Figure 9. nhx1 nhx2 Mutants Are Sensitive to Osmotic Stress.

Wild-type (Col-0) and nhx1 nhx2 mutant lines L2 and L14 were grown

individually in capped test tubes adapted to hydroponic culture in LAK

medium with 1 mM KCl for 3 weeks and then transferred to LAK medium

with 20% PEG6000. The fresh weight of plants (n = 4 per line) was

determined at the indicated times. Single mutants with alleles nhx1-2,

nhx1-1, and nhx2-1, carried in parallel, produced intermediate results

that have been removed for simplicity.

(A) Visual appearance of dehydration symptoms during the assay.

(B) Fresh weight of plants from the onset of stress to completion of the

hyperosmotic treatment. Data are presented as water content relative to

initial values before treatment.

(C) Wild-type and mutant plants recovering from hyperosmotic stress.

Arrows indicate wilted flower buds in the mutants. Note small siliques in

the mutants.


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a reverse genetics approach has shown that the intravacuolar K+

concentration in the Arabidopsis nhx1 nhx2 mutant was only

30% of the wild type and that the vacuolar lumen was more

acidic, presumably due to impaired K+/H+ exchange (Bassil

et al., 2011b). Here, we showed that, indeed, the nhx1 nhx2

mutant had a significant threefold reduction in K+/H+ exchange in

tonoplast vesicles compared with Col-0 plants (Figure 4) and a

marked reduction in the amount of K+ stored in the vacuoles of

leaf mesophyll cells, epidermal cells, and guard cells of stomata

(Figure 7). Using a ratiometric fluorescence assay, Bassil et al.

(2011b) found a reduction in vacuolar K+ in roots of the nhx1 nhx2

mutant. Together, these results indicate that NHX1 and NHX2

proteins account for the majority of the total K+/H+ exchange

capacity in the vacuole of Arabidopsis. The transport activity still

remaining in nhx1 nhx2 plants could be due to the presence of

NHX3 and/or NHX4, which together with NHX1 and NHX2

constitute the class-I family of tonoplast-localized NHX proteins

in Arabidopsis (Yokoi et al., 2002; Pardo et al., 2006). However,

ProNHX3:GUS fusions were preferentially expressed in repro-

ductive organs (Wang et al., 2007); thus, NHX3 is unlikely to

contribute to Na+,K+/H+ exchange in leaves. By contrast,

ProNHX4:GUS expression stained roots and vascular bundles

in leaves (Wang et al., 2007), partially overlapping the expression

Figure 10. Stomatal Conductance under Osmotic Stress.

Wild-type (Col-0) and nhx1 nhx2 mutant line L2 plants were grown in

hydroponic culture in LAK medium with 1 mM KCl for 3 weeks. The rates

of transpiration and photosynthesis were recorded with an infrared gas

analyzer. Leaves were allowed to equilibrate before treatment for at least

10 min and were then subjected to osmotic shock by treatment with 20%

PEG6000 (arrows). Shown is a representative experiment of five repe-

titions with independent plants.

(A) Stomatal conductance.

(B) Photosynthetic rate.

Figure 11. Daily Shifts in Leaf Turgor.

The turgor of leaves of wild-type Col-0 (black traces) and nhx1 nhx2

mutant line L14 (gray traces) growing in hydroponic culture in the

greenhouse was measured with a patch-clamp pressure probe. Note

that leaf turgor pressure and the pressure recorded by the probe are

inversely proportional. The turgor pressure in the leaf patch is opposed to

the magnetic pressure of the clamp, which is kept constant, and the

pressure probemeasures the difference betweenmagnetic pressure and

the relative turgor value. Thus, high pressure values mean lower leaf

turgor pressure. Black and white boxes in the horizontal bars represent

dark and light periods (8/16 h).

(A) Plants growing in 0.1 mM K+ were transferred to 1 mM K+ at the time

indicated by the arrow.

(B) Plants were transferred from 1 to 10 mM K+ (arrow).

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of NHX1 and NHX2. The nonvacuolar, class-II NHX proteins,

NHX5 and NHX6, localize in the Golgi and trans-Golgi network,

where they may regulate endosomal pH (Bassil et al., 2011a).

Other cation/proton exchangers that may add to the residual

Na+,K+/H+ activity in the nhx1 nhx2 plant are Cation/H+ Ex-

changer (CHX) and K+ Efflux Antiporter (KEA) proteins (Pardo

et al., 2006; Chanroj et al., 2012). Arabidopsis CHX proteins (28

members), which are also thought to mediate K+ transport and

pH homeostasis, have been localized to the plasma membrane

and various intracellular compartments, but not to the tonoplast

so far (Chanroj et al., 2012). The KEA proteins (six isoforms in

Arabidopsis) are thought to regulate K+ homeostasis in organ-

elles (Chanroj et al., 2012). Thus, the available evidence strongly

suggests that NHX1 and NHX2, the two most highly expressed

members of the class-I group of NHX proteins, are also the main

players in the active accumulation of K+ in the vacuole. It is worth

noting that disruption of active accumulation of K+ in the vacu-

oles of nhx1 nhx2 plants resulted in greater retention of K+ in the

cytosolic pool (Figure 12). This implies that, while the plasma

membrane potential, which is negative inside the cell, drove the

acquisition of extracellular K+, further transit to the vacuole was

impeded by the tonoplast membrane potential, which is positive

in the lumen relative to the cytoplasm, in the absence of an active

transport mechanism that is capable of accumulating K+ against

its electrochemical gradient. The converse situation was found in

transgenic tomato expressing Arabidopsis NHX1, where en-

hanced recruitment of K+ into vacuoles occurred at the expense

of a diminishing cytosolic pool (Leidi et al., 2010).

Plant cells expand by accumulating solutes, absorbing water,

generating turgor pressure, and extending the cell wall. The

vacuolar K+ pool plays a fundamental biophysical role and, jointly

with other vacuolar osmolytes, drives osmotic changes and

water movements (Leigh, 2001). Plants starved for K+ show,

among other disorders, smaller sizes of aerial parts, decreased

water content, reduced turgor, impaired stomatal regulation, and

reduced transpiration (Mengel et al., 2001; White and Karley,

2010). All these symptoms are linked to the fundamental role that

intracellular K+ plays as osmoticum.Arabidopsismutants lacking

NHX1 and NHX2 were smaller than control plants at all external

K+ regimes, and this was more apparent in shoots than in roots

(Figures 3 and 5). Shoot size was strictly correlated with the

amount of NHX1 and NHX2 activity remaining. Whereas single

mutants had near normal shoot development, leaky double

mutants of the nhx1-1 nhx2-1 genotypewere significantly smaller

than the wild type, and complete knockout plants (nhx1-2

nhx2-1) were severely stunted (see Supplemental Figure 2 on-

line). Supplemental Na+ could partially recover growth of the

nhx1 nhx2 mutant, since nontoxic concentrations of Na+ may

substitute for K+ as osmoticum (Rodrıguez-Navarro, 2000; Bassil

et al., 2011b). Reduced leaf size in nhx1 nhx2 mutant plants

appeared to be a consequence of compromised cell expansion

and not a reduction in the number of cells (Figure 8) (Bassil et al.,

2011b). The relative growth rate of plant cells is a function of the

internal hydrostatic or turgor pressure and the yield threshold

and extensibility of the cell wall. Tissues of the nhx1 nhx2mutant

consistently showed reduced water contents, which correlated

with K+ contents on a dry weight basis. This indicates that the

diminished size of the vacuolar K+ pool hindered water uptake

and that these plants were no longer able to supply the vacuole

with sufficient amounts of K+ for the normal expansion of leaf

cells (Figure 7). Accordingly, leaf turgor pressure measured with

a leaf patch pressure probe was lower in the nhx1 nhx2 mutant

plants than in control Col-0, and this difference increased

steadily over time at low (0.1 to 1 mM) external K+ (Figure 11A).

Transfer to 10 mM K+ stabilized leaf turgor, although daily shifts

in leaf turgor due to light/dark transitions and stomatal function

were dampened in the mutant plants. These findings are coher-

ent with the prevailing view that K+ in the vacuolar pool acts as

the major osmoticum driving water uptake and cell expansion.

Consequently, NHX1 and NHX2, as key players in the creation of

the vacuolar K+ pool, are important determinants of plant growth.

Plants with compromised K+ acquisition also showed reduced

sizes (Hirsch et al., 1998), but phenotypes under nonstarving

conditions were not as dramatic as those found in the nhx1 nhx2

mutant plants.

How Are Na+ Ions Compartmentalized in the Vacuoles

of nhx1 nhx2Mutant Plants?

We have shown that Arabidopsis NHX1 overexpression in to-

mato imparted tolerance to NaCl, which was related to the

preemptive accrual of K+ in vacuoles and improved K+ retention

after stress imposition, but did not enhance the ability to com-

partmentalize toxic Na+ ions into the vacuole (Leidi et al., 2010).

Figure 12. Higher Cytosolic K+ Concentrations in Roots of the nhx1 nhx2

Mutant.

Cytosolic K+ activities (a and b) and plasmamembrane potentials (c and d)

were directly measured by double-barreled K+-selective microelec-

trodes in single epidermal root cells of the wild type (a and c) and nhx1

nhx2 mutant line L2 (b and d). Shown are representative traces of five

replicates.


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Similar findings were obtained by overexpression of the tomato

protein NHX2 (Rodrıguez-Rosales et al., 2008). The long-standing

view is that, owing to the reduced volume of the apoplastic

space, the principal if not the only line of defense of plant cells to

avert cell injury by extracellular salt accumulation is to rely on the

sequestration of salt inside the large central vacuoles. Thereby,

plant cells avert ion toxicity and reduce their osmotic potential to

facilitate water uptake (Oertli, 1968; Flowers et al., 1991; Munns,

2002). The discovery that vacuolar NHX proteins were capable

of exchanging Na+ and H+ across the tonoplast led to the now

widespread view that NHX proteins mediate this critical process

in plants faced with a saline environment (Apse et al., 1999;

Gaxiola et al., 1999; Blumwald, 2000; Quintero et al., 2000).

However, this notion has been recently challenged based on the

biochemistry of NHX proteins, which do not discriminate be-

tween Na+ and K+ or have a preference for K+ transport (Venema

et al., 2002; Rodrıguez-Rosales et al., 2009; Jiang et al., 2010).

The lack of correlative evidence between greater salt tolerance

and the enhancement of Na+ accumulation in different plant

species overexpressing NHX proteins from various sources has

also been pointed out (Rodrıguez-Rosales et al., 2009; Jiang

et al., 2010). Recently, Bassil et al. (2011b) have shown that

NHX1 and NHX2 proteins play a comparatively greater role in K+

homeostasis than in Na+ sequestration. Here, we showed that

mutant plants of genotype nhx1 nhx2 were extraordinarily sen-

sitive tomoderate KCl concentrations (10 to 20mM) but they did

not show greater susceptibility to NaCl compared with the wild

type (Figure 6). In fact, salt-related growth retardation was

proportionally less in the nhx1 nhx2 plants than in thewild type at

50 to 100 mM NaCl (Figure 6; see Supplemental Table 1 online),

and the inclusion of moderate amounts of NaCl in the nutrient

solution containing 20 mM K+ alleviated K+-associated toxicity

symptoms (Bassil et al., 2011b). Notably, nhx1 nhx2 mutant

plants accumulatedmoreNa+ in their shoots than thewild type at

100 mM NaCl (Figure 6; see Supplemental Table 1 online).

Together, these findings raise the questions of how Na+ gets

compartmentalized into the vacuoles of Arabidopsis and which

transport proteins underlie this process (Jiang et al., 2010). It is

now apparent, at least in Arabidopsis, that ion transporters other

than NHX1 and NHX2 mediate the influx of Na+ into the vacuolar

lumen. Biochemical analyses suggest the potential operation of

NHX1 andNHX2 asNa+/H+ antiporters in the tonoplast (Figure 4),

but genetic evidence rules out any significant contribution of

NHX1 and NHX2 in the compartmentation of Na+ (Figure 6). The

budding yeast VNX1 protein, a member of the type II calcium

exchange family, catalyzed Na+/H+ and K+/H+ exchange, but not

Ca2+/H+ exchange, in vacuole-enriched fractions with a Km of

22.4 and 82.2 mM for Na+ and K+, respectively (Cagnac et al.,

2007). Suggestions that members of the calcium/cation antipor-

ter and CHX exchanger superfamilies may also mediate Na+/H+

exchange at the plant tonoplast have not been confirmed ex-

perimentally (Zhao et al., 2009; Chanroj et al., 2012). In this

regard, it is intriguing that genetic inactivation ofNHX1 andNHX2

reduced simultaneously Na+/H+ and K+/H+ exchange capacity in

tonoplast vesicles and that no specific Na+/H+ exchange activity

was unmasked by removing NHX1 and NHX2, while Na+ accu-

mulation still proceeded under salinity stress (Figure 6). Electro-

physiological studies have shown that nonselective SV channels

permeate K+ and Na+ into the vacuolar compartment. Under salt

stress, plant cells accumulate Na+ in the vacuole and release

vacuolar K+ into the cytoplasm. SV channels are thought to

mediate K+ release, but it appears that concomitant Na+ leakage

from the vacuole is impeded as luminal Na+ blocks the SV

channel in Arabidopsis (Ivashikina and Hedrich, 2005). In con-

trast with K+ ions, Na+ could not be released bySV channels even

in the presence of a 150-fold gradient (lumen to cytoplasm). This

property of the SV channel guarantees that K+ can shuttle across

the vacuolar membrane while maintaining the Na+ stored in this

organelle. However, since vacuoles of glycophytic plants may

accumulate up to 80 mM Na+, cytosolic Na+ concentrations

remain at 10 to 30 mM, and the tonoplast membrane potential is

;30 mV, positive in the lumen relative to the cytosol (Carden

et al., 2003; Tester and Davenport, 2003), it appears unlikely that

passive permeation by SV channels would account for mean-

ingful accumulation of Na+ inside vacuoles. In summary, there is

no likely candidate(s) yet to account for the Na+ accumulation

that occurs in the absence of NHX1 and NHX2 in Arabidopsis.

NHX Proteins Facilitate Stomatal Movements

The nhx1 nhx2 mutant exhibited delayed stomatal closure and

thus pronounced leaf turgor loss compared with the wild type.

Regulation of stomatal aperture, preventing excess transpira-

tional vapor loss, relies on turgor changes in two highly differen-

tiated epidermal cells surrounding the pore, the guard cells.

Increased guard cell turgor due to solute accumulation results in

stomatal opening, whereas decreased guard cell turgor following

solute release promotes stomatal closing. The main solutes

involved in the osmoregulation process of guard cell are sucrose

K+, and accompanying anions (malate and chloride), depending

on the environmental conditions. K+ salts are highly mobile and

energetically cheap solutes and, consequently, guard cells accu-

mulate K+ salts in large quantities to open the stomata (MacRo-

bbie, 2006). Arabidopsis CHX20 is a putative K+/H+ exchanger

that appears toplay a role in guard cell osmoregulation throughK+

fluxes and possibly pH modulation (Padmanaban et al., 2007).

However, CHX20 localizes to the endomembrane system and

not to the tonoplast; thus, this protein has been suggested to

associate with vesicles that traffic among various subcellular

membranes.

Scanning electronmicroscopy/EDX data confirmed that guard

cells produced significantly more K+ counts than neighboring

subsidiary cells and mesophyll parenchyma cells (Figure 7), and

this correlated with the expression of NHX1 and NHX2 in guard

cells (Figure 2). Inactivation of both genes reduced by half the K+

content in the cells of leaves, including the guard cells, and

compromised stomata function and development. Guard cells in

mutant plants appeared wrinkled and much thinner than those in

the wild type (Figure 8G), demonstrating that cell expansion is

particularly hindered in the cell type with the most abundant

expression of NHX1 and NHX2. Stomata, however, were not

completely dysfunctional. Wild-type plants responded to an

osmotic challenge in hydroponic culture with a rapid closure of

stomata (Figure 10). Mutant plants nhx1 nhx2 responded more

slowly and reached the reduced transpiration rates of the wild

type only after 90 min of treatment, suggesting that the fast

1138 The Plant Cell

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response mediated by rapid K+ fluxes was impaired in the

mutant. In these experimental conditions, the stomatal conduc-

tance prior to stress imposition was higher in the mutant plants

than in the wild type, suggesting that the closure of stomata is

unconditionally impaired in the nhx1 nhx2 mutant line, perhaps

due to the aberrant development of guard cells (Figure 8G).

Leaves undergo diurnal rhythms of day/night changes in turgor

pressure that are a sensitive indicator of stomatal dynamics and

plant water status. The pressure probe monitors the leaf turgor

response that is directly associated with stomatal movements

(Ache et al., 2010). In low K+, the nhx1 nhx2 line displayed daily

rhythms of stomatal closure and opening that indicated some

preservation of regulated stomatal function, but nonetheless the

leaf turgor pressure decreased steadily over several days as the

plants wilted (Figure 11). Supplementation with 10 mM K+ had

the opposite effect and partially stabilized turgor in the leaf of

nhx1 nhx2 mutants over a few days, while dampening the daily

turgor shifts that remained unaffected in the wild-type plants.

These observations are further evidence of dysfunctional sto-

matal regulation in nhx1 nhx2 mutant plants that is presumably

linked to the reduced ability to compartmentalize K+ in the

vacuoles of guard cells (Figure 7). Delayed stomatal closure and

loss of leaf turgor was observed in Arabidopsis mutants lacking

the SnRK protein kinase OPEN STOMATA1 and the SLOW

EFFLUX ANION CHANNEL-ASSOCIATED1 (Ache et al., 2010).

These proteins trigger membrane depolarization in guard cells in

response to abscisic acid, followed by K+ release and stomata

closing. By analogy with electrophysiological measurements

demonstrating similar plasma membrane potentials in root cells

of the wild type and nhx1 nhx2 mutant (Figure 12), the plasma

membrane potential in guard cells is expected to remain at

normal values even though the cytosolic K+ concentration might

be higher in the nhx1 nhx2 plants.

In summary, NHX1 and NHX2 are vacuolar K+/H+ exchangers

essential for active K+ uptake at the tonoplast, osmotic adjust-

ment, and turgor regulation, and they play a unique role in

stomata function.

METHODS

Growth Conditions and Physiological Determinations

Seeds of Arabidopsis thaliana ecotype Col-0 and the nhx mutant lines

were stratified for 2 to 4 d at 48C and then germinated at room temper-

ature in plastic holders containing mineral wool imbibed in distilled water.

Seedlings were transferred to 8-liter plastic containers for hydroponic

culture. Amodified Long Ashtonmineral solution (Hewitt, 1966) with 1mM

K+ and nominally free of Na+ and NH4+ (LAK medium) was used as base

solution for hydroponic cultures. The final composition of the LAK base

solution was as follows: 1mMKH2PO4, 2mMCa(NO3)2, 1mMMgSO4, 30

mM H3BO3, 10 mM MnSO4, 1 mM ZnSO4, 1 mM CuSO4, 0.03 mM

(NH4)6Mo7O24, and 100 mM Fe2+ as Sequestrene 138-Fe, pH ;5.3. In

experiments with higher K+ levels (10 and 20mM), supplemental K+ was

added as K2SO4. For low (<1mM) K+medium, KH2PO4 was replaced by

NaH2PO4 and K+ was added as K2SO4. For salinity treatments, Na+ was

added as NaCl to the LAK medium. Hydroponic pots were incubated in

a controlled growth chamber with the day/night regime of 25/206 28C,

40/60% relative humidity, 14/10 h illumination, and 250 mmol m22 s21

PAR.

Tomeasure Na+ and K+ contents, salt treatments were administered to

2-week-old plants in hydroponic culture. For salinity treatments, NaCl

was added at 25mM increments every 12 h until reaching 50 and 100mM

Na+ in the nutrient solution. Typically, treatments were performed for two

additional weeks before sampling unless specified otherwise. For ion

content analyses, plants were separated into shoots and roots, and fresh

weight was measured. Roots were washed thoroughly in tap water and

blotted dry before weighing. Dry weight was also measured after drying

samples at 708C for 48 h in a forced-air oven to obtain water contents

(grams of water per gram of dry weight). Na+ and K+ were extracted by

autoclaving finely ground material and measured by atomic absorption

spectrophotometry (Perkin-Elmer 1100B). For scanning electron micros-

copy/EDX analysis, pieces of freshly harvested, fully expanded leaves

were processed as described previously (Leidi et al., 2010). Elemental

analysis was performed focusing the excitation x-ray beam into the

vacuoles exposed in fractured cells. The number and energy of the x-rays

reemitted from the specimen were measured by an energy-dispersive

spectrometer, and counts pertaining to C, O, K, Na, Ca, and Mg atoms

were recorded. Results are presented as the percent of K+ and Na+

counts relative to total counts. Plant extracts obtained from the remaining

parts of samples were used to measure ion concentrations by atomic

absorption spectrophotometry. All statistical analyses were performed

with the software package SPSS version 19.

Leaf gas exchange was determined using the open gas exchange

system Li-6400 (LI-COR) equipped with the chamber head (Li-6400-40;

LI-COR) that allowed full control of light, CO2, and humidity. Stomatal

conductance (gs; mmol m22 s21) and the net photosynthetic rate (AN;

mmol m22 s21) were measured in 3-week-old plants of the wild type and

mutant line L2 grown hydroponically in LAK standard solution. Leaf

responses to osmotic shock were recorded in fully expanded leaves,

attached to the plant, under ambient CO2 and a saturating PPFD of 150

mmolm22 s21. Leaveswere allowed to equilibrate under those conditions

for at least 10min and then subjected to osmotic shock by 20%PEG6000

in LAKmedium. Measurements were recorded every 30 s over a period of

120min. A total of 12measurements for each genotype (six plants per line

and two measurements per plant) were recorded.

Leaf turgor was monitored every 5 min by means of ZIM-patch clamp

pressure probes (ZIM Plant Technology) attached to the youngest fully

expanded leaves. This system allows noninvasive continuous recording

of leaf turgor bymeans of an inversely and highly correlated value, the leaf

patch clamp pressure (Zimmermann et al., 2008; Ache et al., 2010).

Estimates of leaf water potential and osmotic potential were performed

with a dew point microvoltmeter (HR 33T) and a C-52 sample chamber

(Wescor) as described (Wullschleger and Oosterhuis, 1986). Leaf water

contents were measured either by estimates of relative water contents

using leaf discs (RWC = [fresh weight 2 dry weight/rehydrated weight 2

dry weight] * 100) or by measuring total tissue water content (water

content = [leaf fresh weight – leaf dry weight]/leaf dry weight). Leaf area

was determined by the program D-Scan (Scan Analytics) on real-size

images of rosette leaves.

Genetic Methods

Mutant lines with T-DNA insertions were obtained from the SALK collec-

tion (SALK Institute). Alleles and SALK lines used in this work were

SALK_065623 (nhx1-2), SALK_034001 (nhx1-1), and SALK_036114

(nhx2-1) (Alonso et al., 2003). Alleles are named as described by Bassil

et al. (2011b). Insertionmutant information was obtained from the SIGnAL

website (http://signal.salk.edu) and confirmed experimentally. Positions

of T-DNA insertion sites are shown in Supplemental Figure 1A online.

Mutant nhx1-2 has a T-DNA insertion at nucleotide +1246 relative to the

start codon, whereas mutant nhx2-1 carries the insertion at nucleotide

+2429. The T-DNA insertion in line SALK_034001 occurred at the


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59-untranslated region (nucleotide 2114, allele nhx1-1) of NHX1. Homo-

zygous mutant lines were identified by kanamycin resistance selection

combined with diagnostic PCR screening with allele-specific primers

designed to amplify wild-type or mutated loci.

To testmutant complementation inArabidopsis, plasmid pGreen-AtNHX2

was constructed by subcloning the complete NHX2 coding region in XbaI-

BamHI restriction sites of the vector pGreenII-35S-nos-Hyg (John Innes

Centre; http://www.pgreen.ac.uk). The construct pGreenAtNHX2 was co-

electroporated with pSoup vector into the Agrobacterium tumefaciens

GV3101 strain, and the resulting bacterial clones were used to transform

the nhx1-1 nhx2-1 Arabidopsis double mutant by the floral dipping method

(Clough and Bent, 1998). Hygromycin-resistant T1 transgenic plants were

selected on Murashige and Skoog agar medium supplemented with 20 mg/

mL hygromycin B. Complementation tests were performed in LAK medium

supplemented with a range of KCl concentrations.

For yeast mutant complementation and protein purification, a hexa-His

tag was introduced at the 39 end of the NHX2 cDNA by PCR using the

primers 59-GCCAGGATCCTCAGTGATGGTGATGGTGATGCGATCCAC-

GAGGTTTACTAAGATCATGGCTGC-39 and 59-CACTCGAGGAAAGAT-

GACAATGTTCGC-39 and the complete open reading frame of NHX2 as

template. The resulting in-frame fusion was sequenced and introduced

into the XhoI-BamHI sites of plasmid pDR195, generating plasmid

pDR195-NHX2:His6. The tagged protein NHX2:His6 was checked for

complementation of the budding yeast (Saccharomyces cerevisiae) strain

AXT3 (Dena1-4::HIS3, Dnha1::LEU2, and Dnhx1::TRP1) (Quintero et al.,

2000) by the drop-test method using plates of Arg phosphate medium

supplemented with 70 mM NaCl and yeast peptone dextrose medium

supplemented with 30 mg/L hygromycin B.

Biochemical Methods

Isolation of tonoplast vesicles from rosette leaves and measurement of

cation/proton exchange were as described (Barkla et al., 1999), except

that fluorescence quenching of 9-amino-6-chloro-2-methoxy-acridine

wasused tomonitor the formation anddissipationof pHgradients. Purified

tonoplast vesicles (50mg of protein) were added to a buffer containing 250

mM mannitol, 10 mM 1,3-bis[tris(hydroxymethyl)-methylamino]propane–

MES, pH 8.0, 100 mM tetramethyl ammonium chloride, 3 mM MgSO4,

and 1 mM 9-amino-6-chloro-2-methoxy-acridine (1 mL final volume).

Fluorescence was recorded with a Hitachi fluorescence spectrophotom-

eter (FL-2500) in a thermostated cell (268C) at excitation and emission

wavelengths of 415 and 485 nm, respectively (slit-width, 10 nm). The

reaction mixture was stirred and maintained at 268C throughout the

transport assays. Formation of acid-inside pH gradients were startedwith

the addition of 1.5 mM ATP–1,3-bis[tris(hydroxymethyl)-methylamino]

propane, pH 8.0. When fluorescence stabilized, the initial rate of dissi-

pation was measured after the addition of NaCl or KCl salts. Exchange

rates are expressed as fluorescence recovery relative to fluorescence

values prior to ATP addition (DF/Fmax), per minute and milligram of

protein. Exchange rates were fitted to Michaelis-Menten kinetics using

nonlinear regression with KALEIDAGRAPH (Synergy Software).

His-taggedArabidopsisNHX2waspurified fromyeast cellsbyNi2+-affinity

chromatography and inserted into artificial vesicles of soybean phospho-

lipids as described (Venema et al., 2002). Measurement of NHX activity of

His-tagged NHX2 was conducted in pyranine-loaded proteoliposomes as

previously described for NHX1 (Venema et al., 2002). Proton efflux coupled

to cation influx was monitored from the increase of pyranine fluorescence

(463-nm excitation; 510-nm emission).

Histochemical Methods and Microscopy

A genomic DNA fragment upstream of the ATG start codon of NHX2

encompassing ;3.1 kb of the promoter region was fused with the GUS

reporter gene in plasmid pBI101, and the resulting construct was intro-

duced into wild-type Arabidopsis Col-0 plants. Three independent trans-

genic lines were assayed for GUS expression and they showed coherent

results. Stress treatments were applied to seedlings by a 5-h incubation in

160 mMNaCl, 40 mM LiCl, or 320 mM sorbitol in half-strength Murashige

and Skoog medium (Yokoi et al., 2002). Histochemical GUS analyses

were performed as described (Jefferson et al., 1987). The GUS expres-

sion pattern was determined in young seedlings and organs of mature

plants at different stages of development, including roots, rosette leaves,

young flowers, and green siliques. Samples were incubated at 378C for 18

h in GUS staining buffer (200 mM phosphate buffer, pH 7.2, 0.1% [v/v]

Triton X-100, and 1 mMX-Gluc) and then placed for 5 h in 90% ethanol to

remove chlorophyll. Photographs of GUS-stained tissues were taken

using a Zeiss Axioskop microscope equipped with Nomarski optics and

the Zeiss AxioVision software.

Electrophysiological Measurements

For electrophysiological experiments, Arabidopsis (Col-0 and the nhx1-2

nhx2-1 mutant line) was grown aseptically and vertically for 15 d on solid

LAK medium with 1 mM K+ and 0.9% (w/v) Phytagel. Cytosolic K+

activities in epidermal root cells were directly measured by double-

barreled K+-selective microelectrodes. Seedlings were mounted in plex-

iglass chambers (volume of ;1.1 mL), and continuous perfusion of the

assay solution containing 2 mM CaCl2 and 0.1 mM KCl, pH 6.0 (10 mM

MES-BisTris propane) was maintained at a constant flux rate of;10 mL

min21. Microelectrode pretreatment and backfilling were as described

previously (Leidi et al., 2010). The microelectrodes were filled with a K+-

sensor cocktail containing K+ ionophore I (cocktail B, cat. No. 60398;

Fluka, now part of Sigma-Aldrich) dissolved in a mixture of polyvinyl-

chloride/tetrahydrofuran (40 mg mL21) at a ratio of 30:70 (v/v) (Mithofer

et al., 2005). Impalements were performed in epidermal root cells at 5mm

from the apex. The signals from the K+-selective and voltage barrels were

recorded and simultaneously subtracted by a high-impedance differential

amplifier (FD223; World Precision Instruments). The difference was

calibrated before and after experiments with different KCl solutions

(from 1 to 500 mM KCl), maintained at a constant ionic strength by the

addition of MgCl2 (Walker et al., 1995). Calibration curves showed slopes

of around 49 mV/pK+. The impalements were stable for at least 20 min.

Accession Numbers

Sequence data from this article can be found in the Arabidopsis Genome

Initiative or GenBank/EMBL databases under the following accession

numbers: NHX1 (At5g27150), NHX2 (At3g05030), NHX3 (At3g06370),

NHX4 (At5g55470), NHX5 (At1g54370), and NHX6 (At1g79610). Notation

of NHX genes is as described by Yokoi et al. (2002).

Supplemental Data

The following materials are available in the online version of this article.

Supplemental Figure 1. Molecular Genetics Analyses of nhx1 and

nhx2 Mutants.

Supplemental Figure 2. Growth of Double Knockout Mutant nhx1-2

nhx2-1 in Different Potassium Regimes and upon Complementation

with NHX2.

Supplemental Table 1. Shoot Growth and Water Content, and K+

and Na+ Concentrations in Shoots and Roots of Col-0 and nhx1-1

nhx2-1 Mutant Plants.

Supplemental Table 2. Leaf Area, Specific Leaf Weight, and Succu-

lence of Col-0 and nhx1-1 nhx2-1 Mutant Plants.

1140 The Plant Cell

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ACKNOWLEDGMENTS

We thank Francisco J. Quintero and Miguel A. Botella for helpful

suggestions, Antonio Diaz Espejo for help with leaf gas exchange

measurements, and Imelda Mendoza and Maria A. Parrado for technical

assistance. This work was supported by grants from Ministerio de

Ciencia e Innovacion (cofinanced by the European Regional Develop-

ment Fund) to J.M.P. (BIO2009-08641 and CSD2007-00057) and J.A.F.

(CTM2011-30356). Z.A. was supported by the Junta de Ampliacion de

Estudios–Consejo Superior de Investigaciones Cientificas Fellowship

Program. Maria A. Parrado and A.D.L. were supported by the Programa

Nacional de Potenciacion de Recursos Humanos del Plan Nacional de

Investigacion Cientifica.

AUTHOR CONTRIBUTIONS

V.B., E.O.L., Z.A., A.D.L. and B.C. performed research on the physiolog-

ical, biochemical, and genetic data. L.R. and J.A.F. conducted the

electrophysiological measurements. E.O.L., B.C., and J.M.P. designed

the research and wrote the article.

Received December 26, 2011; revised February 20, 2012; accepted

March 5, 2012; published March 20, 2012.

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ion exchangers nhx1 and nhx2 mediate active potassium ... · ion exchangers nhx1 and nhx2 mediate...

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